The original goal of the federally-funded Human Genome Project had been to complete the sequence of the human genome at ten-fold coverage by the year 2005. With dramatic acceleration in pace, a partial draft has recently been presented. Rather than decreasing the need for rapid, inexpensive DNA sequencing, however, this feat has spurred the need for rapid, inexpensive sequencing of nucleic acids. Completion of the draft human genome sequence has also spurred a need for methods and apparatus for analyzing directly the complex collection of genome-encoded proteins, collectively termed the proteome.
With respect to DNA sequence needs, there is growing interest in sequencing the genomes of non-human organisms, including bacteria, plants and animals.
More importantly, the burgeoning fields of molecular pathology and pharmacogenomics will require the resequencing of multiple genes from individual patients. Molecular pathology relates to the diagnosis, and often formulation of a prognosis, for human diseases by identifying mutations in particular genes. Pharmacogenomics refers to understanding how allelic differences that exist in all human populations affect the therapeutic response, and susceptibility to side effects, of individuals to drugs.
As the need to sequence genes from individual patients grows, so will the demand for point of care sequencing capability. There will need to be a shift from large, centralized, high throughput DNA sequencing facilities that only exist at well-funded academic research centers and genomics companies to small, less complicated, middle-throughput gene sequencing systems that can be installed in the majority of hospitals and clinics. This shift in the market for DNA sequencing technologies will put a premium on reducing the cost of reagents and making the sample processing steps as simple and seamless as possible.
In the late 1970s, Sanger et al. developed an enzymatic chain termination method for DNA sequence analysis that produces a nested set of DNA fragments with a common starting point and random terminations at every nucleotide throughout the sequence. Lloyd Smith, Lee Hood, and others modified the Sanger method to use four fluorescent labels in sequencing reactions enabling single lane separations. This resulted in the creation of the first automated DNA sequencers, which used polyacrylamide slab gels for separations. More recently, fluorescent energy-transfer dyes have been used to make dye sets that enhance signals by 2- to 10-fold and simplify the optical configuration.
Automated fluorescent capillary array electrophoresis (CAE) DNA sequencers appear to be the consensus technology to replace slab gels. Capillary gel electrophoresis speeds up the separation of sequencing products and has the potential to dramatically decrease sample volume requirements. The 96-channel capillary electrophoresis instrument, MegaBACE™, which is commercially available from Amersham Biosciences, Inc. (Sunnyvale, Calif.), uses a laser-induced fluorescence (LIF) confocal fluorescence scanner to detect up to an average of about 625 bases per capillary (Phred 20 window) in 90 minute runs with cycle times of two hours. Confocal spatial filtering results in a higher signal-to-noise ratio because superfluous reflections and fluorescence from surrounding materials are eliminated before signal detection at the photomultiplier tube (PMT). Accordingly, sensitivity at the level of subattomoles per sequencing band is attainable. Confocal imaging is also particularly important in microchip analysis systems using capillary electrophoresis, where the background fluorescence of a glass or plastic microchip may be much higher than that of fused silica capillaries. Capillary array electrophoresis systems will solve many of the initial throughput needs of the genomic community for DNA analysis. However, present methods for low volume sample preparation still present a significant barrier to increased throughput and reduced cost.
While fluorescent DNA sequencers are improving the throughput of DNA sequence acquisition, they have also moved the throughput bottleneck from sequence acquisition back towards sample preparation. In response, rapid methods for preparing sequencing templates and for transposon-facilitated DNA sequencing have been developed, as have magnetic bead capture methods that eliminate centrifugation. Thermophilic Archae DNA polymerases have been screened and genetically engineered to improve fidelity, ensure stability at high temperatures, extend lengths, and alter affinities for dideoxynucleotides and fluorescent analogs. These improvements have resulted in lower reagent costs, simpler sample preparation, higher data accuracy, and increased read lengths.
The sequencing community has also developed higher throughput methods for preparing DNA templates, polymerase chain reaction (PCR) reactions, and DNA sequencing reactions. Sample preparation has been increasingly multiplexed and automated using 96- and 384-well microtiter, multi-channel pipettors, and laboratory robotic workstations. In general, these workstations mimic the manipulations that a technician would perform and have minimum working volumes of about a microliter, although stand-alone multi-channel pipettors are being used to manipulate smaller volumes.
A typical full-scale sample preparation method for DNA shotgun sequencing on capillary systems begins by lysing phage plaques or bacterial colonies to isolate subcloned DNA. Under some circumstances it may be desirable to PCR-amplify the subcloned DNA insert to exponentially increase its concentration in the sample. Next, exonuclease I (ExoI) and arctic shrimp alkaline phosphatase (SAP) are added to perform an enzymatic cleanup reaction to remove primer and excess dNTPs that interfere with cycle sequencing. ExoI is used to degrade the single-stranded primers to dNMPs without digesting double-stranded products. SAP converts dNTPs to dNPs and reduces the dNTP concentration from 200 μM, as used for the PCR reaction, to less than 0.1 μM for use with fluorescent sequencing. The reaction is performed at 37° C. and then heated to 65° C. irreversibly denature the ExoI and SAP.
Because PCR amplification can produce excess template DNA for cycle sequencing, the ExoI/SAP treated PCR sample can be diluted five-fold before cycle sequencing. This reduces the concentration of contaminants into a range that causes less interference with capillary electrophoresis analysis. Cycle sequencing reagents are added, typically with fluorescently labeled dye primers or terminators and the reaction is thermal cycled to drive linear amplification of labeled fragments. Finally, after cycling, the samples are post-processed, typically by ethanol precipitation or spin filtration, resuspended in formamide, another denaturant, or water, and the sample is electrokinetically injected into the capillary electrophoresis system.
This workflow has resulted in a dramatic improvement in the performance of the MegaBACE™ system, and similar work flows currently appear to be the methods of choice for other capillary electrophoresis systems as well. Using actual samples from single plaques and colonies of human genomic random subclones or Expressed Sequence Tags (ESTs), this workflow with linear polyacrylamide as a separation matrix has improved the success rate of samples over 200 base pairs from about 60% to 85–90%, and has improved the average read length from about 400 to greater than 600 bases. Furthermore, this method has proven to be quite robust.
While the above sample preparation methods have greatly increased throughout, the cost of reagents remains a major component of the cost of sequencing. Capillary electrophoresis requires only subattomoles of sample, but presently samples are prepared in the picomole range. Reducing the reaction volume will therefore reduce the cost of DNA sequencing and still provide enough material for analysis. However, substantial reductions in reaction volume can only be achieved if satisfactory methods can be developed for manipulating and reacting samples and reagents. Ideally, such a method would be automated and configured to produce multiple samples at one time. Moreover, it would be desirable to integrate such a method as a module capable of interfacing with additional components, such as capillary electrophoresis and a detector for separation and analysis.
Several devices have been designed to aid in the automation of sample preparation. For example, U.S. Pat. No. 5,720,923 describes a system in which small cycling reactions take place in tubes with diameters as small as 1 mm. The tubes are subsequently exposed to thermal cycles produced by thermal blocks to effect the desired reaction. Multiple samples may be processed in a single tube by drawing in small amounts of sample, each of which are separated in the tube by a liquid which will not combine with the sample. Fluid moves through the tubes by means of a pump. These features are incorporated into a system which automatically cleans the tubes, moves sample trays having sample containing wells, and brings the tubes into contact with the wells in the sample trays.
U.S. Pat. No. 5,785,926 discloses a system for transporting small volumes of sample. In this system, at least one capillary tube is used to transport small amounts of sample. A precision linear actuator connected to a computer controlled motor acts as a pneumatic piston to aliquot and dispense liquid using the tube. The sample amount is monitored by an optical sensor that detects the presence of liquid within the capillary segment. The system includes a fluid station containing liquids to be deposited and a positioning device for positioning the transport capillary.
U.S. Pat. No. 5,897,842 discloses a system for automated sample preparation using thermal cycling. In this system a reaction mixture is pumped into a capillary tube. One end of the tube is sealed using pressure from an associated pump while the other end is sealed by pressing the tube against a barrier. The pump also serves to move fluid within the tube. Once the ends are sealed, the tube is exposed to thermal cycles. In this system a robotic transfer device moves the tubes between the sample preparation station where the pump loads the components of the reaction mixture into the tubes and the thermal cycling station.
In the systems discussed above, it is necessary to first mix together a sample, such as DNA template for sequencing, and reagents, prior to introducing the mixture into a reaction chamber. This intermediate mixing step inevitably requires additional reagent and sample handling steps that results in wastage. For example, if separate micropipets are used to dispense sample and reagent into a mixing chamber, small amounts of sample and reagent will be retained in the respective pipets, and reaction mixture will be retained in the mixing chamber. In a high throughput system the cost of this wastage and providing new or properly cleaned pipets and mixing chambers rapidly mounts. Extent of wastage is often exacerbated by the need to dispense relatively large volumes of liquids containing reaction components at low concentration as a strategy to compensate for inaccuracies in dispensing low volumes of higher concentration liquids. Usually, after the reaction mixture is formed, only a small proportion is required for analysis, and the remainder is discarded.
Thus, there exists a need for means by which a biological sample to be analyzed could be introduced into a reaction chamber without the need to first mix the sample with the reagents necessary to effect the reaction.
U.S. Pat. No. 5,846,727 discloses affinity-capture methods wherein template DNA is immobilized inside a glass capillary tube that serves as a reaction chamber for thermal cycling. The capillary is first prepared by immobilizing biotin molecules to the inner surface of the capillary, followed by charging the column with avidin or streptavidin which binds tightly the biotin. Template DNA to be sequenced is covalently linked to a biotin moiety by PCR, and is then exposed to the avidin inside the capillary. This results in immobilization of the template to the capillary wall through a biotin-avidin-biotin linkage. After unbound template is washed away, sequencing reagent is added, and the contents of the capillary are subjected thermal cycling to activate the sequencing reaction. In this manner it is unnecessary to mix template DNA with sequencing reagent prior to loading the capillary.
However, the method just described requires that biotin be linked to the template DNA by PCR, necessitating setting up and carrying out a reaction even before the sequencing reaction. This requisite preliminary step adds to the time and cost associated with acquiring the sequence data. Furthermore, the immobilization of the DNA is effectively irreversible because the biotin-avidin linkage is so strong it can only be broken using agents that denature avidin, a treatment that would also denature any other protein components in a reaction. As a result the template DNA must stay bound to the inner surface of the capillary. Because the DNA is not free in solution, additional time is required for reaction components to diffuse to the walls where they can interact with the DNA. Furthermore, when it is desired to recycle the capillary, it is necessary to remove the template DNA via denaturation of the avidin, washing and then recharging of the avidin in the capillary, all of which add to time and reagent costs.
Thus, there is continued need in the art for methods to introduce molecules into reaction chambers without an initial sample-reagent mixing step, without the need to attach an affinity capture moiety to all the molecules in the sample, and wherein template immobilization is reversible. In this way reagent costs would be minimized and processing speed maximized.
Capillary array electrophoresis systems and capillary electrophoresis microchip analytical systems can detect subattomoles of DNA sequencing reaction products. This extraordinary sensitivity comes at the cost of reduced tolerance, compared to slab gels, for deviations from the ideal amount of template DNA in the sequencing reactions. For example, if there is too little template DNA in the sequencing reaction, there will be poor yield of fluorescently labeled primer extension products. This results in weak signal strength when the reaction products are scanned by the laser. This prevents the software that analyzes the chromatogram from adequately performing spectral separation, resulting in shorter than average sequence read lengths; the reaction will have to be repeated or the sequence information will be lost.
Too much template DNA causes problems as well, due to overloading of the capillary. While there is adequate yield of fluorescently labeled reaction product, if the template is in excess, it competes with sequencing products for entry into the capillary during electrokinetic injection. The presence of the large template DNA molecules can result in an overall reduction, or sudden drop in capillary current, which can manifest itself in a variety of ways. overloading can cause weak signal strength, late appearance of interpretable fluorescence intensity peaks in the chromatogram, and poor resolution of the reaction products because the fluorescence emission is broad and diffuse. All these effects lead to shorter reads and lower sequencing data quality.
The problem of overloading is typically solved by either diluting the sequencing reaction, or carefully titrating the amount of template DNA introduced into the sequencing reaction. While both these solutions are simple in principle, the former requires repeating the analysis of the reaction, and the latter is difficult to implement using conventional means in a high-throughput system. These means include detecting, and comparing to standard concentration curves, the quantity of fluorescent dye that binds DNA in a sample, or measuring the absorbance of ultraviolet light at 260 nm wavelength, which can be converted into an absolute measure of DNA concentration. Thus, there is continued need in the art for methods to titrate the quantity of template DNA for sequencing reactions to be analyzed using high-throughput capillary electrophoresis systems, where minimizing cost and maximizing speed are crucial.
There is an additional need for an automated system that is able to perform small-scale thermal cycling reactions in a highly parallel manner. The system should allow for rapid preparation of cycling reactions with minimal consumption of reagents. The combination of reducing the amount of reagents required for a reaction and reducing the time required for a reaction will greatly reduce the overall cost of preparation of cycling reactions.
With respect to proteomics, analysis of the proteome requires separation, quantification and identification of large protein collections.
Typically, such analysis is achieved by a combination of different techniques, such as 2-D electrophoresis separation, followed by enzymatic digestion and identification by matrix-assisted laser desorption/ionization mass spectrometry (2D PAGE-MALDI/MS) or by electrospray ionization mass spectrometry (2D PAGE-ESI/MS). Another common approach is LC/LC-MS/MS, i.e., proteins are digested, separated by strong cation exchange liquid chromatography and reversed phase liquid chromatography (LC/LC), and then identified by tandem mass spectrometry (MS/MS). Current limitations include the requirement for extensive sample preparation prior to proteolytic digestion, analyte loss, and low reaction efficiencies at low protein concentrations.
In an alternative, methods and apparatus have been developed that permit both partial purification and mass spectal identification using a single derivatived laser desorption probe. See, e.g., U.S. Pat. Nos. 6,225,047, 6,124,137, 5,719,060. Such methods, however, require specialized equipment and familiarity with mass spectrometers.
There is, therefore, a continued need in the art for an automated system that is able to perform small-scale proteomic reactions in a highly parallel manner. The system should allow for rapid preparation of enzymatic reactions with minimal consumption of reagents. The combination of reducing the amount of reagents required for a reaction and reducing the time required for a reaction will greatly reduce the overall cost of preparation of proteomic reactions while a highly parallel system will improve throughput.